Preparing for recording

Preparing brain tissue for recording is a crucial step. If your tissue is bad, your recordings will be bad. You will find that every lab uses a slightly different protocol. While you could argue that variability in the protocols could make is harder to replicate studies across labs, the most important thing is that you can get stable cells. Variability between protocols is within in reason because there is only so much you can change before you do not get viable tissue. There are papers that claim to have improved protocols but they generally have very little good data to back them up. We cover many differences between protocols and the support or lack of support for these protocols.

Steps to process tissue for recording¶

Generally there are about 3 steps you can take before recording: perfusion, tissue slicing and tissue recovery. The only step that is optional is perfusion, the last two you generally have to do. Most of the variability comes from what solutions you use and how you let the tissue recover.

Perfusion¶

Perfusion is not considered an essential step but many labs perfuse anyways for a variety of reasons. A general definition of perfusion is removal of the blood in the animal by replacing the blood with an osmotically similar fluid. For patch clamp electrophysiology researchers use some version of ACSF. When blood gets into the brain tissue in a living organism is can be quit catostrophic such as during a stroke or TBI. Perfusion does remove blood however, when slicing the tissue any blood is released into the cutting solution and does not necessarily embedded into the tissue. Additionally, there is relatively little blood in the brain. I have noticed in adult animal the blood cells can occasionally form strings that come off the cut blood vessels and get in the way of patching a cells. Generally the blood stays contained within the blood vessels. Another way that way perfusion may improve tissue quality is by preventing oxidative damage and excitotoxicity due to hypoxia Kirmse et al., 2015. Interneurons may be particularly sensitive to oxidative stress.

Chilled vs room temperature perfusion solution¶

For patch clamp electrophysiology you can use a chilled or room temperature solution. The reasoning behind using a chilled solution is that is slows brain metabolism and thus slows degradation of the brain. I have not seen any evidence to support this, but it is a relatively minor consideration since it is easy to chill a solution. One point that several researchers brought up is that if you are going to chill the perfusion solution you should probabably try to keep the brain chilled throughout the extraction and cutting steps. Another thing to point out is that since some rodent brain are relatively small they warm up quickly after extracting the brain thus chilling might provide minimal benefit. Additionally, if the perfusion solution is running through a pump it might warm up by the time it gets to the subject depending on how cold the solution is. The solution will also take time to cool subjects body to a point where the body is not warming up the solution.

Choice of perfusion solution¶

For patch clamp electrophysiology you will want to use some version of ACSF rather PBS. The osmolality of PBS is considerably lower than ACSF. I have seen papers perfuse with standard ACSF or ACSF that has the Na+ substituded with sucrose, choline or NMDG. The reason to use a low Na+ ACSF is that it is thought to prevent excitotoxicity. I have not seen any good evidence that the type of ACSF matters for perfusion but this is hard to test. Some labs use ACSF for perfusion but a Na+ substituted ACSF for cutting so you don’t necessarily need to use the same solution for perfusion and cutting. One thing to note is that choline is a mild cholinergic agonist so if you want to avoid that do not use choline ACSF.

Perfusing the subject¶

To perfuse, you need to sedate the mouse with avertin or isoflurane. You want to avoid anesthetics with long half-life since these could bind to receptors that you are trying to record from. Avertin and isoflurane are fast acting and quick to wear off. Next you need to open the chest cavity and cut the right aorta to allow for the blood to exit the circulatory system. Using a syringe or perstaltic pump you will puncture the right ventrical and inject your fluid of choice. You should see the liver slowly turn form a dark red to a light brown/pink color. For young mice you need to inject about 5-10 mL were as an adult mouse can take up to 25 mL (rats take significantly more) depending on much you want to clear the brain of blood. In adult mice you can inject ~10 mL/min, the younger the mouse the slower you want to go.

Cutting¶

For cutting tissue there are several things to consider: solutions, temperature, cutting apparatus and tissue thickness.

Solutions¶

The same solutions that are used for perfusion can be used for cutting tissue. Typically labs use a ACSF with the Na+ substituted for sucrose, choline or NMDG. There are some methods papers claiming that NMDG is superior but were not really a comprehensive comparison of tissue preparation methods. However, many electrophysiologists will claim that using a low Na+ ACSF is better. Lowering the Na+ is supposed to prevent excitotoxicity. One reason to use a choline or NMDG based cutting solution is that they maintain the ionic balance, since they are cations but, they are to large to travel across the membrane thus preventing excitotoxicity. You will also want to continuously bubble your solution with carbogen while cutting to maintain the pH and O2/CO2 balance. This can be done by just putting a small diameter tube into your cutting well.

To chill or not chill the cutting solution¶

Many labs chill the cutting solution or even turn it into an icy slush because it is supposed to firm up the tissue making it easier to cut and better preserve the tissue. Extremely cold or slushy solutions can cause morphological changes in neurons Kirov et al., 2004Eguchi et al., 2020Fiala et al., 2003. These changes are reversible but, we typically want to minimize any changes to the neurons that happen during the tissue preparation stage. I would recommend chilling the solution since it does seem to prevent the tissue, particularly heavily myelinated sections of the brain, from sticking to the cutting blade.

Cutting apparatus considerations¶

For the cutting apparatus you can use a vibratome or a compresstome. The compresstome requires you to embed the brain in a low melting temperature agarose. For the vibratome it is optional to embed the brain in agarose. On the vibratome you can also put a block of agarose behind the brain to prevent it from moving backwards while cutting. The compresstome compresses the embedded brain while cutting which is supposed to prevent chatter (horizontal lines) and uneven sections that can occur on the vibratome. However, when cutting thick sections on a vibratome chatter is not really an issue. The main issue with the vibratome is making sure the brain is adhered securely to the chuck. I recommend trying both the gel and liquid super glue. I find that the gel is better for brain tissue from young (~P16) mice whereas a more liquid superglue is better for adult mice. You also want to make sure you do not put too much superglue down since it will creep up the brain when immersed in liquid and can impair cutting. Additionally you need to make sure that you wick away most of the liquid around the brain before setting on the superglued on the chuck to prevent the glue from prematurely setting. It always good to compare the types of superglue with your own setup and equipment. For the vibratome or compresstome you use steel safety razor blades. You can also use ceramic or tungsten blades. Ceramic blades (such as Cadence EF-INZ10) can be much sharper than the steel blades thus can help cut through heavily myelinated regions. However, I do not know of any labs using the ceramic blades likely because they are about 5x more expensive. Most vibratomes and the compresstome have a recommended angle to put the blade at. Follow those instructions. For the vibratome I recommend keeping the minimal amount of liquid in the cutting well the prevent the tissue from flapping around. You can use a camel hair brush to float the sections around if you need.

Tissue thickness¶

I have seen tissue cut anywhere from 200 uM to 300 uM. The thicker the tissue the harder it will be to see cells deeper in the tissue. The thicker the tissue the harder it is for oxygen to perfuse into the tissue. Thicker the tissue the more intact cells deeper in the slice will be. The thinner the tissue the more delicate your tissue will be. General rule of thumb is thicker slices for younger mice since there is less extracellular stuff holding the tissue together. In some areas, like the cortex, the dendrites often run parallel to the slice in coronal sections so thicker sections may improve how intact the dendrites are.

Recovery¶

Recovery can consist of one or two parts. Regardless of the cutting method you use you will need to let the tissue recover for at least 30 minutes and often up to an hour before recording. In the two step recovery the sections are removed from the cutting chamber and put into a recovery solution in a heated bath kept at ~32°C. One thing you should consider is that if you are using a recovery solution in a heated bath to keep the sections in <12 minutes. There seems to be some trade off between how good the cells look and how functionally normal the cells are. I have found that prolonged incubation in a choline or NMDG ACSF leads to high access resistance. Additionally, if you are incubating in a low Na+ solution you can also do a Na+ spike in where you gradually increase the Na+ levels (as well as the osmolarity). This will prolong the time you are in the recovery solution. See Ting et al., 2018 for more information on how to use the Na+ spike in.

Solutions¶

The recovery solution is typically the same as the cutting solution. I mostly see labs using choline- or NMDG-ACSF. I have not seen the sucrose-ACSF likely because of the risk of bacterial growth with all the sugar in the solution. If you want to try the Na+ spike in solution see Ting et al., 2018 for the recipe. The Na+ spike is design for use with NMDG but could be adapted to use with choline solution.

After your sections recover in the recovery solution you will transfer your sections to a room temperature ACSF for at least 30 minutes. You can use standard ACSF or a HEPES ACSF. The HEPES helps prevent edema (cell swelling) in the tissue which can be problematic in tissue from adult mice.

Incubation chambers¶

You will need some sort of incubation chamber to hold your sections in. You can buy expensive chambers (AutoMate Scientific), make your own or 3D print your own (I have STL files which you can access here: https://github.com/LarsHenrikNelson/3D_print_neuroscience_tools and there are variety on the web that you can find by just looking).